Indigoid dyes by group e monooxygenases: Mechanism and biocatalysis

Article abstract of DOI:10.1515/hsz-2019-0109

Since ancient times, people have been attracted by dyes and they were a symbol of power. Some of the oldest dyes are indigo and its derivative Tyrian purple, which were extracted from plants and snails, respectively. These 'indigoid dyes' were and still are used for coloration of textiles and as a food additive. Traditional Chinese medicine also knows indigoid dyes as pharmacologically active compounds and several studies support their effects. Further, they are interesting for future technologies like organic electronics. In these cases, especially the indigo derivatives are of interest but unfortunately hardly accessible by chemical synthesis. In recent decades, more and more enzymes have been discovered that are able to produce these indigoid dyes and therefore have gained attention from the scientific community. In this study, group E monooxygenases (styrene monooxygenase and indole monooxygenase) were used for the selective oxygenation of indole (derivatives). It was possible for the first time to show that the product of the enzymatic reaction is an epoxide. Further, we synthesized and extracted indigoid dyes and could show that there is only minor by-product formation (e.g. indirubin or isoindigo). Thus, group E monooxygenase can be an alternative biocatalyst for the biosynthesis of indigoid dyes.

Full text of DOI:10.1515/hsz-2019-0109

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Biological Chemistry ’Just Accepted’ paper
ISSN (online) 1437-4315
DOI: 10.1515/hsz-2019-0109
Research Article
Indigoid dyes by group E monooxygenases:
mechanism and biocatalysis
1
1
1
1,2,
Thomas Heine , Carolin Großmann , Sarah Hofmann and Dirk Tischler *
1
Institute of Biosciences, Environmental Microbiology, TU Bergakademie Freiberg, Leipziger
Str. 29, D-09599 Freiberg, Germany
2
Microbial Biotechnology, Ruhr University Bochum, Universitätsstr. 150, D-44780 Bochum,
Germany
*
Corresponding author
e-mail: dirk.tischler@rub.de
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BioIndigo by group E monooxygenases
Abstract
Since ancient times, people are attracted by dyes and they were a symbol of power. Some of
the oldest dyes are indigo and its derivative Tyrian purple, which were extracted from plants
and snails, respectively. These “indigoid dyes” were and are still used for coloration of
textiles and as a food additive. The traditional Chinese medicine also knows indigoid dyes as
pharmacologically active compounds and several studies support their effect. Further, they are
interesting for future technologies like organic electronics. In these cases, especially the
indigo derivatives are of interest but unfortunately hardly accessible by chemical synthesis. In
recent decades, more and more enzymes have been discovered that are able to produce these
indigoid dyes and therefore gained attention from the scientific community. In this study,
group E monooxygenases (styrene monooxygenase and indole monooxygenase) were used for
the selective oxygenation of indole (derivatives). It was possible for the first time to show that
the product of the enzymatic reaction is an epoxide. Further, we synthesized and extracted
indigoid dyes and could show that there is only minor by-product formation (e.g. indirubin or
isoindigo). Thus, group E monooxygenase can be an alternative biocatalyst for the
biosynthesis of indigoid dyes.
Keywords: dye production; epoxidation; flavoprotein; indole monooxygenase; styrene
monooxygenase.
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Introduction
One of the oldest pigments used by humans is Tyrian purple. It was and is one of the most
valuable dyes as a gram still costs about 3000 €. Tyrian purple, which is chemically 6,6`-
dibromoindigo, is a derivative of the well-known denim colorant indigo. Substituted indigo
derivatives are termed as indigoid dyes (Figure 1). In recent years, these indigoid dyes gained
interest in several other application areas, as they represent interesting candidates for
pharmaceutical, food and semiconductor industries (Ma et al., 2018b). In particular, indigoid
dyes have been shown to function as antibiotic (Namgung et al., 2019) and against several
types of cancer as they have antineoplastic or antileukemic activity (Xiao et al., 2002; Dua
et al., 2014). Furthermore, they have hemostatic, antipyretic, anti-inflammatory, and sedative
properties (Dua et al., 2014). Thus, they are among others of interest for the treatment of
allergies, Alzheimer’s disease and delayed hypersensitivity (Kapadia et al., 1998; Wu et al.,
2
005; Hössel 1999; Hoessel et al., 1999; Guengerich et al., 2004a,b; Eisenbrand et al., 2004;
Leclerc et al., 2001; Kunikata et al., 2000; Lin et al., 2012; Dua et al., 2014; McClay et al.,
005; Farias-Silva et al., 2007; Pathak and Madamwar 2010; Xiao et al., 2002). A recent
2
study showed that encapsulation of indigoid dyes into inorganic hosts can be useful for the
production of hybrid pigments and photo-functional materials (Petermayer and Dube, 2018).
Moreover, they can be used in dye-containing polymers, photoswitches, on memory chips and
in high-performance organic semiconductors (Petermayer and Dube, 2018; Fleischmann
et al., 2015; Harrer, 2012; Uehara et al., 1987; Irimia-Vladu et al., 2012; He et al., 2014;
Głowacki et al., 2012). In the latter case, especially the high-cost derivative Tyrian purple is
of interest. Thus, it is of importance to produce indigoid molecules selectively in order to have
pure compounds for the subsequent applications.
While indigo itself is already produced at industrial scale (about 50 000 tonnes per year), the
indigo derivatives are hardly accessible by chemical means. Therefore, bacterial sources for
the production of indigoid dyes have been exploited for several years and the biosynthesis
routes of indigoid dyes can be seen as one alternative of aerobic indole detoxification. The
other alternative is the degradation of indole to use it as a source for carbon and energy (Arora
et al., 2015; Ma et al., 2018b).
Indole and its derivatives are widely distributed in the environment as they occur naturally in
alkaloids, phytohormones, plant flower oils, pigments, fossil organic matter, signaling
molecule and product of the tryptophan metabolism but also anthropogenically as product of
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BioIndigo by group E monooxygenases
pharmaceutical industries (Lee et al., 2015; Arora et al., 2015; Sharma et al., 2010; Ma et al.,
2
018b). Due to that extensive exposure, a variety of microorganisms was found to be able to
degrade and transform indole and in the course of that, many enzymes (monooxygenases and
dioxygenases) were identified that enable the formation of indigoid dyes (reviewed in Heine
et al., 2018a; Ma et al., 2018b). These often initiate the degradation of indole by mono- or
dioxygenation at the double bond of the pyrrole moiety or at the benzene moiety (Heine et al.,
2018a). Intermediates of these reactions that were often detected are indoxyl,
dihydroxyindole, indole-2,3-dihydroindole and oxindole leading to isatin and/or anthranilate
as central intermediates (Fukuoka et al., 2015; Yuan, 2011; Sadauskas et al., 2017; Fujioka
and Wada, 1968; Ma et al., 2018b).
Despite the fact that many oxidoreductases were identified that are able to oxidize indole, for
most of them it seems to be not the natural substrate (Ma et al., 2018b). However, recently
identified monooxygenases (IndA, IifC) were found in Acinetobacter, Burkholderia and
Cupriavidus strains located in a gene cluster that enables indole degradation in these bacteria
(
Sadauskas et al., 2017; Lin et al., 2015; Qu et al., 2017; Zhang et al., 2018; Ma et al.,
018a). This and an associated anthranilate degradation cluster has already been identified by
Tischler et al., in 2012 and was proposed to be involved in aromatic compound degradation
Tischler et al., 2012). The initiating monooxygenases were originally assigned as styrene
2
(
monooxygenase (group E monooxygenase, GEM) due to sequence homology. GEMs are
flavin adenine dinucleotide (FAD) dependent enzymes, which require for a nicotinamide
adenine dinucleotide (NADH) dependent FAD reductase as the second component for the
supply of reduced cosubstrate (Montersino et al., 2011). However, in accordance with the
recent findings, GEMs have been divided into two subgroups: styrene monooxygenase (SMO)
and indole monooxygenase (IMO) (Heine et al., 2018b). Finally, the monooxygenases IndA
and IifC can now be assigned to the latter group of IMOs where indole is supposed to be the
natural substrate.
However, it is to date not solved, which product is actually formed by IMOs: indole oxide or
directly indole-2,3-dihydrodiol (Boyd et al., 1997; O'Connor et al., 1997; Sadauskas et al.,
2017). In this study, we established a method published by Dupard-Julien et al., 2007 to
clarify this issue. Therefore, we selected four GEMs that were initially characterized by Heine
et al. (2018a): AbIndA is known to be part of a gene cluster that allows for indole
biodegradation. VpIndA1 is located in a gene cluster also containing the respective indole
degradation genes and in addition a fusion protein of the IMO. It was proven a promising
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sulfoxidase (Tischler et al., 2018). RoIndA1 is part of a similar gene cluster while one gene
that is supposed to be required for indole degradation is disrupted (Tischler et al., 2012).
RoIndA1 is one of the best-studied IMOs (Tischler et al., 2010; Riedel et al., 2015; Paul
et al., 2015). Additionally, the SMO GpStyA was included as it showed one of the highest
activities of all GEMs in preliminary tests.
These GEMs were used for the production of indigoid dyes, which were identified by Vis-
spectral and MS analysis.
Results
Biotransformation of Indole Derivatives to Indigoid Dyes
The four monooxygenases have been successfully produced recombinantly in Escherichia
coli as described previously (Tischler et al., 2009, 2018; Riedel et al., 2015). During gene
expression, the cells turned dark blue, which is an indication of indigo formation. This
happens due to tryptophanase (TnaA) activity of host cells leading to significant amounts of
indole from tryptophan (Li and Young, 2013), which is subsequently oxygenated by the
present recombinant monooxygenases. It indicated a successful enzyme production. The
-1
proteins were obtained at high purity (yield of 3 to 38 mg enzyme l , depending on type and
culture medium) and were subsequently assayed for styrene epoxidation activity. They
-1
-1
showed activities as follows; GpStyA (570 ± 90 mU mg ) > AbIndA (139 ± 16 mU mg ) >
-1
-1
RoIndA1 (121 ± 22 mU mg ) ≥ VpIndA1 (121 ± 9 mU mg ). In all cases the (S)-enantiomer
was formed at high selectivity (> 94% ee) (Heine et al., 2018a). These rates and
enantioselectivity are in a comparable range to a number of previously investigated styrene
-1
epoxidizing enzymes (range 26 to 2100 mU mg ) (Otto et al., 2004; van Hellemond et al.,
007; Tischler et al., 2010, 2018; Toda et al., 2012; Tan et al., 2019; Gursky et al., 2010;
2
Riedel et al., 2015).
As group E monooxygenases are two-component flavoprotein monooxygenases, they rely on
a support of reduced FAD in order to perform catalysis. To circumvent additional enzymes
and cofactor limitation, the assays were performed with 1-Benzyl-1,4-dihydronicotinamide
(
BNAH) as direct FAD reductant (Paul et al., 2015). For each enzyme, a concentration-
dependent performance was determined (Figure 2). It turned out that all four enzymes behave
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similarly with respect to BNAH supply with an optimal concentration between 8 and 12 mM.
For the other experiments, the BNAH concentration was set to 10 mM.
Having those active monooxygenases and proper support of reduced FAD for catalysis, an
assay for indole oxygenation was set up as described in the materials and methods section.
The following substrates, indole, 6-fluoroindole, 6-chloroindole and 6-bromoindole were
applied and the formation of respective indigoid dyes was followed spectrophotometrically
(
Figure 3). Herein it has to be mentioned, that the monooxygenases reach between 6 and 80%
activity with indole compared to styrene: VpIndA1 (80%) > AbIndA (60%) > RoIndA1 (17%)
GpStyA (6%). Except for RoIndA1, unsubstituted indole is the best substrate for all
monooxygenases (Figure 3). The halogenated derivatives reach an activity of only about 20 to
0% if compared to indole. Interestingly, while 6-fluoroindole is not a good substrate for
>
6
RoIndA1, the chlorinated and brominated analogs work as good as the unsubstituted indole.
The extinction coefficients that either had been obtained from literature or that were
determined from the upscaling experiments (Figure 4 and 5) were employed to calculate
kinetic parameters. However, as discussed above this might not provide actual access on the
monooxygenase functionality as several catalytic steps occur in the reaction course of indole
to indigo. This is true for the derivatives as well. Despite this remark, there was still some
variance determined from the kinetic experiments, which might be due to a different substrate
acceptance of enzymes or due to a different dimerization velocity of generated indoxyl
derivatives, respectively.
In order to produce more dye from the three different halogenated substrates upscaling
experiments were performed with VpIndA1 and GpStyA. We obtained an amount of 2.3 to
5
.5 mg halogenated indigoid dyes as well as 1.3 to 5.4 mg dimethoxy-substituted indigoid
dyes (Table 1). This indicates that the conversion is better if the halogen substituent is bigger
and/or has a lower electronegativity and that substitution is beneficial at the 5-position
compared to 6- and 7-substituted indoles. However, this might be different depending on the
respective enzyme.
An FT-ICR-MS analysis was done to identify the generated dyes. It should be mentioned, that
this method does not discriminate between the indigo and the structural isomers indirubin and
isoindigo. However, indirubin is much more stable as indigo in organic solvents and should
get visible after a certain time (Meyer, 2002). We investigated that for several
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biotechnologically produced dyes but did neither detect a corresponding change in color nor
an absorbance shift to lower wavelengths in DMSO (Perpète et al., 2006). Thereof, we
hypothesize that mainly indigo (derivatives) are formed by GEMs, what remains to be proven
by additional analysis.
However, Table S2 summarizes the results of the mass-spectrometry assignment. The
observed mass-to-charge ratio fits the calculated ratios for the expected products.
Tracing epoxide formation by GEMs
As mentioned above, the intermediate that is formed by the monooxygenase IndA is yet
unknown due to instability. Although several hypotheses about this intermediate were
published, we presumed an epoxide to be formed by IMOs, as GEMs are generally known to
act as epoxidases on double bonds. To clarify this, we attempted to trap the epoxide
intermediate by derivatization with N,N-diethyldithiocarbamate (DTC) by an adapted method
of Dupard-Julien et al., 2007 (see materials and methods section). The expected derivatization
mechanism is displayed in Scheme 1.
In advance to an enzymatic conversion, we performed control experiments to exclude
unspecific reactions with other possible products or interference with components of the
reaction mixture (see supplemental material section 1 and 2). Only the analogical indene
oxide was derivatized by DTC treatment demonstrating the general functionality of the
method (Figure 6A). For the enzymatic assays, we initially conducted the derivatization at
7
0°C after the ending of the reaction. This attempt was successful and an additional peak was
observed eluting about 5 min later than the substrate indole (Figure 6B). The peak area
increased slightly after a biotransformation time up to 10 min. However, parts of the product
continued to react as the formation of indigo was still observed. The derivatization was shown
to be temperature dependent (Dupard-Julien et al., 2007). Nevertheless, to lower the
derivatization temperature to 30°C was also appropriate and allowed to meet the enzymatic
reaction conditions (supplementary material, section 1). Therewith, it was feasible to add
DTC directly to the enzymatic reaction and to trap the epoxide immediately after formation
by the IMO.
The identified product peak was collected after RP-HPLC and subjected to FT-ICR-MS
analysis. The observed mass-to-charge ratio matches the expected ratio for the proposed
hydroxyindole derivative (Table S1, Scheme 1).
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Discussion
In this study, we selected four GEMs that have been characterized previously (RoIndA1,
accession number: ACR43973; VpIndA1, ADU39063; AbIndA, CAG69430 and GpStyA,
GAB22407). All of them perform selectively oxygenations on styrene and indole
(
-derivatives). Styrene and indole degradation are also the two physiological roles described
for GEMs so far (Heine et al., 2018b). However, analysis of the genomic neighborhood of
GEMs indicated, that further functions in aromatic compound degradation are possible.
SMOs are often situated in a styrene degradation cluster allowing for conversion of styrene
into phenylacetaldehyde as central intermediate (Tischler et al., 2012; Heine et al., 2018c).
For several other SMOs, the physiological role is unclear, e.g. for GpStyA which has no
genes in the surrounding that are related to aromatic compound degradation. In contrast, a
highly similar SMO from Prauserella rugosa NRRL B-2295 (WP_030532027) is located
close to a phenol hydroxylase and subsequent enzymes for the attack at the aromatic ring.
Thereof, it might be possible that styrene is degraded in this strain by oxidation at the vinyl
side chain as well as at the aromatic ring system.
The second group of GEMs, IMOs, was recently classified as an independent subgroup and
these monooxygenases are proposed to be involved in indole degradation (Sadauskas et al.,
2017; Heine et al., 2018b) (Figure 7A). The core gene clusters that can be deduced from the
genomic analysis, in which IMO-like enzymes were found, are illustrated in Figure 7B. The
representative cluster (cluster 1) was identified in Acinetobacter and Cupriavidus. It contains
a monooxygenase (IndA, IifC), an associated reductase (IndB, IifD), a short chain
dehydrogenase (SDR) family protein (IndC, IifB) and a dienelactone hydrolase (DLH) family
protein (IndD, IifA). The resulting enzyme cascade was described to constitute the upper
indole degradation pathway to anthranilate as central intermediate (Gröning et al., 2014;
Sadauskas et al., 2017; Zhang et al., 2018; Lin et al., 2015; Qu et al., 2017) (Figure 7A).
IndA catalyzes the oxygenation of indole to an unstable intermediate (vide infra) that can
rapidly decompose to indoxyl, which dimerizes to indigo under aerobic conditions (O'Connor
et al., 1997; Sadauskas et al., 2017). However, the indigo formation is prevented whenever
IndC is present. Then, IndC immediately takes the unstable intermediate, and forms
3
-hydoxyindolin-2-one, which is further converted by IndD to anthranilic acid (Sadauskas
et al., 2017). Genes for the lower degradation pathway from anthranilate to intermediates of
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the tricarboxylic acid cycle were often found for representatives of cluster 1 in proximity or
elsewhere on the genome (Table 2) (Zhang et al., 2018).
Interestingly, we did not find anthranilate degradation genes in strains that harbor clusters 2 to
10 (Figure 7B). Indeed, genes related to the degradation of indolepyruvate and xanthine are
adjacent (Table 2). Further, only cluster 1, 2 and 6 contain a complete set of genes to fulfill
the described degradation route. The IMOs of Variovorax paradoxus EPS belong to the latter
one. Herein, two IMOs are encoded whereby one is a fusion protein between the
monooxygenase and a reductase part (IndA2B) (Tischler et al., 2018). This special
characteristic is known since 2009 when it was discovered and described for the first time for
Rhodococcus opacus 1CP (Figure 7B, Cluster 7) (Tischler et al., 2009). Both clusters
(
6 and 7) contain an SDR-like and a DLH-like protein, although the DLH is disrupted in
strain 1CP. The fusion protein is supposed to act mainly as reductase component and the
associated IndA1 enzymes also possess a much higher specific oxygenation activity (Tischler
et al., 2018; Tischler et al., 2010). Just recently, a novel IMO has been identified in
Burkholderia sp. IDO3, which is able to degrade indole (Cluster 10). It forms a separate
branch from other IMOs as well as SMOs (Ma et al., 2015, 2018a). Interestingly, only strains
where the IMO is arranged in clusters like 1 and 10 have been proven capable to use indole as
a source of carbon and energy (Table 2).
The fourth enzyme of this study is an SMO originating from Gordonia polyisoprenivorans
NBRC 16320 (GpStyA). Although classified as SMO, it is not part of a styrene degradation
cluster and the physiological role is so far unclear. However, it has been shown to provide one
of the highest epoxidation rates of GEMs and therefore was included in this study
(unpublished).
Indole is widespread in the environment and many industrial products. The metabolic fate of
indole has attracted greater interest in the last years, as many significant roles in prokaryotic
but also eukaryotic organisms have been reported (Ma et al., 2018b; Lee et al., 2015; Kim and
Park, 2015). In this study, we used IMOs for the biotransformation of indole to indigoid dyes.
So far, only indole-2-monooxygenases (EC 1.14.13.137) are known to use indole as a
physiological substrate for the biosynthesis of benzoxazinoids (Glawischnig et al., 1999).
Thus, IMOs are the first representatives reported to degrade indole as a physiological
substrate. However, so far only strains that harbor IMO clusters like 1 and 10 have been
reported to grow on indole (Table 2). Conscious that also the gene clusters for the degradation
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of the intermediated anthranilate are missing in many cases, it has to be proven whether there
are further routes for indole degradation or further roles of IMOs in that organisms. For
example, the degradation of indolepyruvate or xanthine or the biosynthesis of other
compounds.
All of the tested IMOs were able to produce indigoid dyes from indole precursors. We also
tested styrene as a reference substrate. For all of the enzymes, the activity with indole is lower
compared to styrene (Figure 3). For GpStyA this corresponds to only about 6% of residual
activity. This correlates with the fact that this enzyme is denoted as styrene epoxidizing
monooxygenase and indole is not the physiological substrate.
It was possible to convert a variety of indole derivatives with GEMs that contain different
substituents at various positions. In addition, the production of indigoid dyes with mixed
substituents at both indole moieties is possible. This illustrates the broad applicability and
product range that can be produced with GEMs. However, substituents at positions 2 and 3 of
the indole heterocycle are problematic for the enzyme, what is reasonable, as this is the
probable oxidation site.
Several hypotheses exist about the formed product. A possible route via an indole oxide and
subsequent chemical or enzymatic hydrolysis to yield an indole-2,3-dihydrodiol followed by
dehydration to yield indoxyl is discussed in literature (Boyd et al., 1997; O'Connor et al.,
1
997; Sadauskas et al., 2017). Otherwise, the indole oxide can undergo an intramolecular
rearrangement known as arene oxide-NIH shift reaction (Cerniglia et al., 1984). Furthermore,
styrene oxide isomerases catalyze such an intramolecular rearrangement and it was found that
these enzymes are beneficial for indigo production employing styrene monooxygenases as
initial catalysts (O'Connor et al., 1997; Miyamoto et al., 2007). However, the indigo
formation does also happen in the absence of styrene oxide isomerase, thus two mechanisms
might be possible in dependence of the environment. In case of dioxygenases, it was shown
that indole gets di-oxygenated to the cis-indole-2,3-dihydrodiol which spontaneously
dehydrates to indoxyl (Ensley et al., 1983; Mermod et al., 1986; Murdock et al., 1993).
In this study, we were able to do a qualitative analysis of the formed product, an indole-2,3-
oxide, for two different IMOs (AbIndA and VpIndA1). Therewith, we propose that GEMs
convert indole to an unstable epoxide, which rapidly hydrolyzes to a diol (Scheme 2). This
diol can rapidly decompose to indoxyl, which spontaneously dimerizes to indigo. However, as
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BioIndigo by group E monooxygenases
soon as other enzymes grab the epoxide or the diol, further degradation to anthranilic acid is
observed (Sadauskas et al., 2017).
Conclusion
Indigoid dyes are interesting compounds for various applications. Enzyme cascades can be
used to produce indigo derivatives in an eco-friendly manner (Namgung et al., 2019).
Together with novel strategies for the generation of water-soluble forms that make them
easier to handle (Hsu et al., 2018), these indigo derivatives are better accessible for industrial
applications. GEMs are candidates for the production of these dyes and in particular, IMOs, as
they use indole as a physiological substrate. In addition, we were able to prove for the first
time that an IMO epoxidizes indole as an initial step of the indole degradation pathway.
Materials and methods
Chemicals, strains, gene expression, and protein purification
Styrene and indole(-derivatives) were purchased from Sigma (Steinheim, Germany), TCI
(
Eschborn, Germany), Carl Roth (Karlsruhe, Germany), ABCR (Karlsruhe, Germany),
Fluorochem (Hadfield, Derbyshire, UK), Alfa Aaser (Karlsruhe, Germany).
E. coli DH5α was used for cloning purposes and E. coli Bl21 (DE3) pLysS was used for gene
expression (Sambrook and Russell, 2001). RoIndA1 was obtained by PCR amplification and
VpIndA1, AbIndA and GpStyA were ordered by gene synthesis as described previously
(
Tischler et al., 2010, 2018; Riedel et al., 2015). The codon usage of the latter constructs was
adapted to that of Acinetobacter baylyi ADP1 (Riedel et al., 2015). All monooxygenase genes
were subcloned into a pET16bP vector system for expression.
Gene expression was done as described previously by induction with isopropyl-β-D-
thiogalactopyranosid (Tischler et al., 2010; Riedel et al., 2015) or by using Terrific broth
(
TB) autoinduction medium (Studier, 2005). Successful protein synthesis was apparent by a
blue color formation resulting due to the indigo production. The target enzymes were purified
by immobilized metal ion chromatography as published earlier (Tischler et al., 2010; Riedel
et al., 2015).
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BioIndigo by group E monooxygenases
Enzyme assays and biotransformation of indole (-derivatives)
Initial activity determination was done by the standard group E monooxygenase assay with
styrene as substrate (Tischler et al., 2010; Riedel et al., 2015). For all assays, NADH as an
electron donor was replaced by the mimic 1-benzyl-1,4-dihydronicotinamide (BNAH) (Paul
et al., 2015). The ideal BNAH addition was determined by means of separate standard assays
with varying BNAH concentrations between 3 and 14 mM.
Additionally, indole(-derivatives) were supplied as substrate by means of the standard
monooxygenase assay but in a 96 well plate. Formation of respective indigoid dyes was
monitored spectrophotometrically (SpectraMax M2e; SoftMax Pro; Molecular Devices,
USA). Therefore, UV/Vis-spectra (330-730 nm) of each indigoid dye was monitored over 2 h
at 10 min intervals. Maximum absorption of each product was plotted over time to determine
the specific activity.
Tracing of the indole-epoxide formation
Formation of epoxides during GEM conversion of indole was done according to the protocol
of Dupard-Julien et al. (2007). After the enzymatic reaction, the epoxide is derivatized with
N,N-diethyldithiocarbamate (DTC; 30°C or 70°C for 20 min) and unreacted DTC is
3 4
eliminated by acidification with H PO . In a second attempt, we added DTC directly to the
assay to extract the epoxide immediately from the reaction mixture and prevent its potential
decomposition. Indole, 1-indanol, isatin, indene, and 1-phenyl-1,2-ethanediol served as
controls but should not be derivatized by the method. The enzymatic reaction was set up
according to the standard monooxygenase assay as described in the previous section.
Therefore, AbIndA was applied as biocatalyst and indole was used as substrate. Due to the
instability of indole-2,3-oxide, we used indene-1,2-oxide as a reference compound. The
derivatives were applied to RP-HPLC analysis using a Eurospher C18 column (125 mm
length × 4 mm i.d., 5 mm particle size, 100 Å pore size; Knauer, Berlin, Germany) with a 40
-1
to 75% acetonitrile gradient over 10 min at a flow rate of 1 ml min (25°C). Eluting
substances were monitored with a UV/Vis-detector. The detected product peak containing the
generated derivative was collected, concentrated and subjected to mass spectrometry.
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Upscaling, indigo purification, and spectral analysis
Upscaling of dye production was done by means of 25 ml assays for halogenated and
methoxy-substituted indoles. The assays contained the following components: glycerol (5%
-1
v/v), Tris-HCl buffer (20 mM pH 7.5), dithiothreitol (1 mM), catalase (20.8 U ml ), FAD
50 µM), BNAH (10 mM) and the respective substrate (2 mM). The mixed components were
(
tempered to 30°C and the reaction was started by addition of enzyme (2.8 mg). After 1 h
reaction at 30°C (120 rpm), the produced indigoid dye was pelleted by centrifugation
(
2
11 000 g). The supernatant was removed; the pellet was washed twice with 2 ml dH O and
dried at 60°C for 22 h. The product was weighed and partially dissolved in DMSO. Non-
dissolved dye was removed from the solution by centrifugation and extinction was measured
at the respective absorption maximum. DMSO was removed at 60°C under vacuum and the
resulting pellet was weighed. The amount of indigoid dye that was dissolved in DMSO was
calculated by the mass difference before and after dissolution experiments. The extinction
coefficient in DMSO was calculated for each indigoid dye according to Lambert-Beers law
(
E = ε • c • d). The generated indigoid dyes were subjected to mass spectrometry.
Mass spectrometry
Fourier-transform ion cyclotron resonance mass spectrometry (FT-ICR-MS) experiments
were conducted on a 15 T solariX FT-ICR-MS from Bruker Daltonics, equipped with an LDI
source and operated in negative ion mode, with a scan range from 46.06 to 1000.00 Da.
Resulting data sets had a size of 8 M and 16 scans were accumulated for each analysis. The
sample was prepared directly by applying 1.5 µl of the undiluted sample solution per spot on
a stainless steel target. For LDI analyses a Smartbeam II laser (frequency tripled Nd:Y laser,
λ = 355 nm, 3 ns pulse duration, 500 µJ pulse energy) in the integrated LDI source was used.
Furthermore, a medium focus, 20 or 250 laser shots per ICR fill and a fixed laser power of
4
0% or 70% were used for the sample analysis of indigoid dyes and the DTC-derivative,
respectively.
Peak picking, calibration, and molecular formula assignment were accomplished using Bruker
Daltonics software DataAnalysis 5.0 (SR1).
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BioIndigo by group E monooxygenases
Acknowledgments
This research was funded by the Saxon Government (SAB) grant number 100263733 and the
DECHEMA (Max-Buchner Fellowship to D.T.) grant number MBFSt 3339. We thank Jan
Zuber from the Institute for Analytical Chemistry and Lucas Siegel (TU Bergakademie
Freiberg) for support in the implementation of the analytics.
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BioIndigo by group E monooxygenases
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Tables and figures
Table 1
Cell-free production of indigoid dyes by GEMs. The extinction coefficient of
the produced dyes was determined in DMSO.
-1
-1
Product
Indigo
Enyzme Yield [mg] Yield [%]
λ
max [nm]
620
Extinction coefficientDMSO [L mol cm ]
VpIndA1
2.2
2.3
3.4
5.5
5.4
4.1
1.3
4.3
33.6
30.8
41.1
52.4
67.0
50.9
16.1
50.4
3660
4271
3112
2506
1992
5709
13 036
1809
6
,6'-Difluoroindigo
590
GpStyA
6
,6'-Dichloroindigo
605
6
,6'-Dibromoindigo
610
5
6
7
,5'-Dimethoxyindigo
,6'-Dimethoxyindigo
,7'-Dimethoxyindigo
665
580
VpIndA1
620
6
-Bromoindigo
610
Table 2
Assignment of IMO-related gene clusters and adjacent genomic regions.
Anthranilate degradation
Adjacent Gene (Clusters)
C
Strain
cluster on the genome
Acinetobacter baylyi ADP1*
Acinetobacter sp. O153
Acinetobacter sp. JW
Acinetobacter baumannii ATCC19606
Burkholderia sp. H160
Cupriavidus sp. SHE
Anthranilate degradation
Anthranilate degradation
Anthranilate degradation
Anthranilate degradation
-
-
-
Indolepyruvate degradation
Anthranilate/catechol/phenol degradation
Anthranilate/catechol/phenol degradation
Yes
Yes
Yes
Yes
Yes
Yes
Yes
Yes
Yes
Yes
1
Cupriavidus necator H16
Halomonas halocynthiae DSM14573
Marinobacterium stanieri S30
Pseudomonas psychrotolerans NS337
2
3
Delftia acidovorans SPH-1
-
No
Gemmobacter nectariphilus DSM 15620 Indolepyruvate degradation
Thermocrispum agreste DSM 44070
Sphingobacterium lactis DSM 22361
No
No
No
Xanthine degradation
Indolepyruvate degradation
4
5
Mesorhizobium sp. F7
-
No
Gypsy moth gut metagenome
Deep-sea sediment metagenome
-
-
-
Xanthine degradation
6
7
Variovorax paradoxus EPS*
Rhodococcus opacus 1CP*
-
-
No
No
Nocardia farcinica IFM 10152
Paenarthrobacter aurescens TC1
-
No
No
8
9
Dehydrogenase/Cyclase
Gordonia sp. I37
Streptomyces auratus AGR0001
-
No
No
Xanthine degradation
1
0
Burkholderia sp. IDO3
-
No
C – Cluster. Strains that are proven to degrade indole are underlined. *IMOs used in this study.
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Figure 1
(Left) Structure of the dye indigo shaded with the respective color. (Right)
Indigoid dyes are derivatives of indigo that can differ in color and chemical properties.
Middle) The most prominent representative is Tyrian purple, which owns two bromide
(
substituents at position 6 and 6’. The figure shows this valuable dye produced in this study
smoothed over a sheet of paper.
Figure 2
Optimization of BNAH concentration for the monooxygenase assay and
biotransformation. The reaction was followed the standard assay with varying concentrations
of the electron donor 1-benzyl-1,4-dihydronicotinamide (BNAH). The optimal BNAH
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BioIndigo by group E monooxygenases
concentration for the monooxygenases is located between 8 and 12 mM (blue box). The
chemical structure of BNAH is indicated.
Figure 3
Several halogenated indole derivatives were applied as substrates for GEMs.
The shown relative activities refer to the determined activity with indole, which is indicated
above the respective bars.
Figure 4
mg of Tyrian purple were obtained from biotransformation, extraction and vacuum drying
left) and it was dissolved to a great extent in dimethyl sulfoxide (right) prior spectral
analysis.
The product of 6-Bromoindole conversion by GpStyA: Tyrian purple. About
5
(
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BioIndigo by group E monooxygenases
Figure 5
UV/Vis-spectral analysis of the indigoid dyes dissolved in DMSO. The
purified indigoid pigments were employed for the determination of extinction coefficients
Table 1).
(
Figure 6
RP-HPLC chromatograms showing DTC derivatization of (A) the control
indene oxide and (B) an AbIndA conversion of indole. Reference assays without DTC are
shaded in grey while derivatization assays with DTC are shaded in red.
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Figure 7
(A) Proposed degradation pathway for indole. The product of the
monooxygenase is so far nor known. However, should be transformed by the activity of IndC
to 3-Hydroxyindolin-2-one and by IndD to the central intermediate anthranilate (Sadauskas
et al., 2017).
(
B) Comparison of IMO-related gene clusters. Representative strains for each core gene
cluster are listed in Table 2 together with associated aromatic compound degradation genes.
The designation of the genes was adapted according to the succession in the indole
degradation pathway, where the monooxygenase is the initiating enzyme (see also Qu et al.,
2017; Heine et al., 2018 b). Styrene degradation-related gene clusters can be found in Heine
et al., (2018c).
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BioIndigo by group E monooxygenases
Scheme 1
Illustration of the indole derivatization with DTC following the method of
Dupard-Julien et al., (2007). Herein, indole is epoxidized by a GEM to indole 2,3-oxide. This
epoxide can now react with DTC to form stable esters that can be separated and identified by
mass spectrometry. Unreacted DTC can be decomposed by acidification.
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BioIndigo by group E monooxygenases
Scheme 2
The predicted reaction mechanism for the formation of indigo by GEMs.
Indole is epoxidized to form indole-2,3-oxide which undergoes chemical or enzymatic
hydrolysis to yield an indole-2,3-dihydrodiol. This is subsequently dehydrated to yield
indoxyl that spontaneously dimerizes to indigo (A). When the enzymes IndC and IndD are
present, the diol is further degraded to anthranilic acid via 3-hydroxyindolin-2-one (B)
(
Sadauskas et al., 2017). The tinting of the enzyme shapes is according to Figure 7.
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Supplementary Material
Section 1
Derivatization of epoxides in a imo biotransformation.
Figure S1
(Top) Chromatogram of the reaction products of a 30 min biotransformation of
indole by AbIndA without derivatization. The substrate indole elutes after 5.27 min.
(Middle) The nicotinamide cofactor elutes after 4.02 min.
(Bottom) DTC (N,N-diethyldithiocarbamate) elutes after 7.40 min.
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Figure S2
Chromatogram of the reaction products of a 5 to 20 min biotransformation of
indole by AbIndA after derivatization with DTC. Herein, DTC derivatization was done after
the respective timepoints. The derivatized product elutes after 10.85 min.
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Figure S3
Chromatogram of the reaction products of a 5 to 20 min biotransformation of
indole by AbIndA after derivatization with DTC. Herein, DTC was added directly to the
reaction mixture to extract the formed epoxide directly after the enzymatic formation. The
derivatized product elutes after 10.84 min.
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